Skip to main content
NCBI home page
NIHPA Author Manuscripts logoLink to NIHPA Author Manuscripts
. Author manuscript; available in PMC: 2014 Aug 4.
Published in final edited form as: Cell Rep. 2013 Jun 13;3(6):1815–1823. doi: 10.1016/j.celrep.2013.05.014

Choroid plexus-derived Otx2 homeoprotein constrains adult cortical plasticity

Julien Spatazza 1,5, Henry HC Lee 2,5, Ariel A Di Nardo 1, Lorenzo Tibaldi 3, Alain Joliot 3, Takao K Hensch 2,4,*, Alain Prochiantz 1,*
PMCID: PMC4119931  NIHMSID: NIHMS606809  PMID: 23770240

Summary

Brain plasticity is often restricted to critical periods in early life. Here, we show that a key regulator of this process in visual cortex, Otx2 homeoprotein, is synthesized and secreted globally from the choroid plexus. Consequently, Otx2 is maintained in selected GABA cells unexpectedly throughout the mature forebrain. Genetic disruption of choroid-expressedOtx2 impacts these distant circuits and in primary visual cortex reopens binocular plasticity to restore vision in amblyopic mice. The potential to regulate adult cortical plasticity through the choroid plexus underscores the importance of this structure in brain physiology and offers novel therapeutic approaches to recovery from neurodevelopmental disorders more broadly.

Introduction

Experience shapes neural circuitry during developmental critical periods (Hensch, 2004). Thus, cognitive ability is weakened enduringly by adverse environments (Nelson et al., 2007), motor maps reflect early musical training (Elbert et al., 1995), native speech sounds sculpt phonemic perception (Kuhl et al., 1992), and sight strengthens the acuity and connectivity of the two eyes in the primary visual cortex. Imbalanced input (“lazy eye”) during this period results in a permanent loss of vision for that eye (Maurer and Hensch, 2012; Prusky and Douglas, 2003), a pathological condition known as amblyopia affecting 2-4% of the human population. As for many neurodevelopmental disorders, there is currently no cure for amblyopia in adulthood (Bavelier et al., 2010).

Cortical inhibition triggers the onset of the critical period (CP) of plasticity for binocular vision around postnatal day (P)21 in mice (Hensch, 2005). Gain- and loss-of-function experiments demonstrate that Otx2 homeoprotein in turn controls the maturational state of a particular subclass of GABA-ergic interneuron containing parvalbumin (PV-cells) (Sugiyama et al., 2008). In Otx2 heterozygous mice, the visual CP does not open, while conversely providing Otx2 protein to wild type PV-cells before P18 opens plasticity ahead of time and anticipates its closure (Sugiyama et al., 2008). The persistent maintenance of Otx2 protein within PV-cells throughout life subsequently restricts plasticity levels in adulthood (Beurdeley et al., 2012). Thus, CP timing can be accelerated, delayed or extended through Otx2 manipulation. Yet, this homeoprotein is neither produced nor confined to postnatal visual cortex, raising the question as to its origin.

While the Otx2 gene is found in several areas of the developing brain, expression in most regions markedly decreases during early ontogenesis (Nothias et al., 1998; Puelles et al., 2004; Rath et al., 2007). A transition from general CNS morphogen to a more narrowly defined role in the pineal gland and retina is suggested (Koike et al., 2007; Rath et al., 2006; Simeone et al., 1993). Indeed following injection into the eye, exogenous Otx2 is found in visual cortical PV-cells and disrupting intraocular synthesis or its translocation into PV-cells impacts visual plasticity (Beurdeley et al., 2012; Sugiyama et al., 2008). But, these findings do not rule out other possible sources of Otx2, during and beyond development. In this study, we demonstrate that the choroid plexus is a global source of Otx2 (Johansson et al., 2013). As a result, knocking-down Otx2 in this structure impairs its transfer into distant PV-cell targets and thus allows the reactivation of plasticity in adult visual cortex. Moreover, a much broader role for Otx2 beyond the visual cortex is suggested.

Results

Otx2 expression in the adult brain and choroid plexus

After eye opening, Otx2 homeoprotein is transferred into the primary visual cortex by an activity-dependent mechanism (Sugiyama et al., 2008), thus triggering PV-cell maturation and CP opening. Given the widespread distribution of PV-cells throughout the forebrain and the presence on their surface of peri-neuronal nets (PNN) required for Otx2 internalization (Beurdeley et al., 2012; Miyata et al., 2012), we examined whether Otx2 translocation might be a common principle, not solely limited to the visual cortex. We have previously shown, by RT-PCR and with the Otx2GFP/+ knock-in mouse line, that the Otx2 locus is silent in the visual cortex (Sugiyama et al., 2008). We now confirmed by in situ hybridization that Otx2 transcripts are not found anywhere throughout the adult cortex (Fig. 1A-D), while evident in dorsal thalamus (dLGN) and superior colliculus as expected (Fig. 1B2, C2). Interestingly, immunohistochemistry (Fig. 1E-L) revealed Otx2 protein across prefrontal, auditory, somatosensory and visual cortices as well as limbic structures (basolateral amygdala BLA, hippocampal CA1).

Figure 1.

Figure 1

Open in a new tab

Non-cell autonomous Otx2 protein is localized throughout the adult cortex.

(A-D) In situ hybridization of Otx2 mRNA performed on adult wild-type mouse brain coronal sections show locus activity in the dLGN, SC and choroid plexus (arrow). Framed regions are magnified and labeled (A1-A2, B1-B2, C1-C2 and D1-D2).

(E-L) Corresponding immunostaining for Otx2 and PV in the regions highlighted in panels A-D reveal Otx2 protein where the Otx2 locus is inactive (scale bar: 100 μm).

(M-N) Quantification of Otx2 co-localization in PV-cells and vice versa (M) and of Otx2 immunostaining intensity per cell (N) across brain regions (3 mice, 5 sections each). Error bars, SEM.

Abbreviations: A1, auditory cortex; BLA, basolateral amygdala; CA1, field of hippocampus; Cg1, cingulate cortex; Cx, cortex; dLGN, dorsal lateral geniculate nucleus; S1, somatosensory cortex; SC, superior colliculus; V1, visual cortex.

In all cortical regions where the protein was present but the locus inactive, the vast majority of Otx2-stained cells also expressed PV (Fig. 1M). In contrast, in the dLGN and superior colliculus where the Otx2 locus is active, Otx2 was not detected in PV-expressing cells (Fig. 1M). In the dLGN, we could not quantify co-localization as PV was present in the neuropil but not in cell bodies. The mean intensity of Otx2 staining per cell was similar in all regions independent of whether Otx2 activity was cell autonomous or not, even though a slightly higher expression may be seen in the cell autonomous regions (Fig.1N).

One potential explanation for this widespread distribution of Otx2 could be a global source from which it is secreted. The choroid plexus, an established site of Otx2 expression (Johansson et al., 2013; Simeone et al., 1993), is a very attractive candidate, known for its intense secretory activity and cerebrospinal fluid (CSF) production (Johanson et al., 2008). The CSF also carries a wide range of signaling molecules acting at the level of the parenchyma. In early development, the choroid plexus provides a niche of proliferative factors (IGF1, FGF2, Shh, retinoic acid) (Alonso et al., 2011; Huang et al., 2010; Lehtinen et al., 2011; Martin et al., 2006) and concentration gradients of guidance molecules for migrating neural adult progenitor cells (Sawamoto et al., 2006). Thus, a similar paracrine release of Otx2 may likewise affect postnatal plasticity by regulating neocortical PV-cells.

In situ hybridization on adult brain sections confirmed the expression of Otx2 in the choroid plexus of the lateral and fourth ventricles (Fig. 2A and insets A1-2), as well as third ventricle (not shown). We next applied a secretion assay developed for other homeoproteins (Engrailed, Pax6) (Di Lullo et al., 2011; Wizenmann et al., 2009) to choroid plexus dissected from the fourth ventricle of adult mice. The assay based on the biotinylation of cell surface proteins (Fig. 2B) demonstrates that around 4.5% of total Otx2 is accessible to the extracellular biotinylation reagent. Accessibility was also true for TrkB, a transmembrane protein, but not for RhoA, a control intracellular protein. Similar results were obtained following the cell surface biotinylation of choroid epithelial cells in culture (not shown).

Figure 2.

Figure 2

Open in a new tab

Otx2 synthesis, secretion and genetic deletion in the choroid plexus.

(A) Sagittal brain in situ hybridization reveals Otx2 expression in adult SC and choroid plexus. Insets show lateral ventricle (A1) and fourth ventricle (A2).

(B) Detection of choroid plexus cell surface proteins with surface biotinylation assay. Inputs show extracellular (TrkB) and intracellular (RhoA) protein. Upon biotinylation of adult choroid plexus, a pool of Otx2 (#5% of total) is found to be extracellular (compare TrkB and RhoA). Note that 2.5% of total extracts and 50% of biotinylated fractions were loaded.

(C) Western blots of mouse choroid plexus and CSF protein extracts probed for: TrkB, a transmembrane protein; TTR, a protein secreted by the choroid plexus; and Otx2. Only Otx2 and TTR are detected in CSF. Film exposure times for mCSF TrkB and Otx2 boxes are 60-fold longer than for mCSF TTR.

(D-G) X-Gal staining of adult ROSA26R mouse brain sections 5 days after single icv injection of vehicle solution (D) or CRE-Tat protein (E-G) reveals recombination in choroid plexus (E, arrow in F). Framed regions are enlarged (D1, E1, F1 and G1).

(H-I) Recombination in Otx2flox/flox mice, 14 days after single icv injection of vehicle solution (-) or CRE-Tat protein (+). (H) Genomic DNA extracted from choroid plexus, retina, dLGN and SC was analyzed by PCR using primers surrounding the flox cassettes (Fossat et al., 2006) yielding an expected 451bp amplicon upon recombination of the Otx2 locus. Only the choroid plexus of CRE-Tat treated animals exhibit partial recombination. (I) Quantification of Otx2 mRNA and protein content in choroid plexus, retina, dLGN and SC of vehicle-treated and CRE-Tat-treated animals. Error bars, SEM (Mann-Whitney U test; 8 mice per group; * p<0.05).

Abbreviations: CSF, cerebral spinal fluid; dLGN, dorsal lateral geniculate nucleus; SC, superior colliculus.

We further examined the presence of Otx2 in the CSF. Given its low protein concentration and limited availability in adult mice, CSF was collected from 10 adult mice. Transtyrethin (TTR), the major thyroid hormone transporter in the CNS, is specifically expressed by the choroid plexus and secreted into the CSF where it represents ∼25% of all proteins (Weisner and Roethig, 1983). Fig. 2C reveals the presence of both Otx2 and TTR in the CSF. In contrast TrkB, also expressed in the choroid plexus (Timmusk et al., 1995), is not detected in the CSF sample (Fig. 2C). In Figure 2C, the exposure time for Otx2 and TrkB in the mouse CSF (mCSF) is 60 fold that of the other panels. The absence of membrane-bound protein, TrkB, in these conditions makes it unlikely that Otx2 in the CSF reflects cellular contamination, but rather secretion from the choroid plexus. However, it also indicates that Otx2 concentration in the CSF is very scarce and that the protein following its release from the cell surface is likely rapidly transported to the brain parenchyma and from there into PV-cells (see discussion).

Knocking-down Otx2 in the choroid plexus decreases its cortical content

To probe the putative impact of Otx2 release into the CSF upon cortical function, we developed a loss-of-function technique. A cell-permeable recombinase (CRE-Tat), obtained by the fusion of the bacterial enzyme with a peptide vector, was injected into the lateral ventricles of adult reporter mice (ROSA26R)(Soriano, 1999). The efficiency of recombination tested five days after a single injection was very high, as demonstrated by the substantial LacZ expression in the choroid plexus (Fig. 2D-F). This access to the choroid plexus was extremely specific as illustrated by the absence of recombination in the dLGN and Superior Colliculus (Fig. 2F-G). The specificity of CRE-Tat targeting is best explained by the CSF clearance activity of the choroid plexus through rapid apical reabsorption (Johanson et al., 2008).

We next turned to Otx2flox/flox mice (Fossat et al., 2006; Nishida et al., 2003) and waited for 14 days after CRE-Tat injection before assessing the efficiency of Otx2 deletion at the level of the genome, mRNA and protein content. Even though the genomic rearrangement in the choroid plexus was not complete, it was specific and did not affect the retina, dLGN or Superior Colliculus (Fig. 2H). Accordingly, a strong and specific decrease in both Otx2 mRNA and protein content (Fig. 2I) was observed in the choroid plexus of CRE-Tat treated animals when compared to vehicle-treated animals (mRNA: Vehicle, 1.00 ± 0.056 VS CRE-Tat, 0.36 ± 0.056, p=0.022 / Protein: Vehicle, 1.00 ± 0.065 VS CRE-Tat, 0.24 ± 0.049, p=0.028), while these parameters were not significantly different from control in the retina, dLGN and SC of CRE-Tat treated animals.

If Otx2 is maintained in adult brain by the choroid plexus, then protein levels in distant cortex should be modified by our CRE-mediated conditional deletion strategy. Indeed, a significant reduction in the number of Otx2-positive cells and their mean fluorescence intensity was observed in the binocular visual cortex (V1b) of CRE-Tat treated animals compared to vehicle (Fig. 3A, F, K, L; cell number: 68.1 ± 3.7%, p = 0.002; mean intensity per cell: Vehicle, 100 ± 1.2 vs CRE-Tat, 82.9 ± 1.2, p = 0.0008). Consistent with our previous work indicating an Otx2 requirement for PV expression and PNN assembly early in life (Sugiyama et al., 2008), a parallel decrease of PV (Fig. 3B, G, K, L; cell number: 74.1 ± 4.7%, p=0.009; mean intensity per cell: Vehicle, 100 ± 1.9 vs CRE-Tat, 91.3 ± 2.0, p=0.0013) and PNN expression (Fig. 3C, H, K; WFA-positive cell number: 68.5 ± 4.2%, p = 0.002) was observed following CRE-Tat injection.

Figure 3.

Figure 3

Open in a new tab

Choroid Otx2 disruption selectively impacts PV-cells in visual cortex.

(A-J) Immunostaining for Otx2, PV, WFA, calretinin (CR) and GABA in layers II-IV of binocular visual cortex (V1b) from adult Otx2flox/flox mice, 14 days after single icv injection of vehicle (A-E) or CRE-Tat protein (F-J). Scale bar: 100 μm.

(K) Percentage of Otx2-, PV-, WFA-, CR- and GABA-positive cells (CRE-Tat versus vehicle) in layers II-IV of V1b. Dashed line, vehicle-treated levels.

(L) Quantification of the mean intensity of Otx2, PV, CR and GABA staining per cell in supragranular layers of V1b (t-test, 4 mice per group; ** p<0.01, *** p<0.001; 700×450 μm area).

(M) Quantification of Western blots of non-cell autonomous Otx2 protein after vehicle or CRE-Tat injection in adult Otx2flox/flox mice (normalized with tubulin, mice 3-4 months old; t-test; 9 mice per group; ** p<0.01).

(N) Quantification of Western blots of non-cell autonomous Otx2 protein in visual cortex during postnatal development (normalized with tubulin; t-test, 5 WT mice for each age; ** p < 0.01).

(O) Quantification of Otx2 mRNA content in the choroid plexus during postnatal development (relative to GAPDH, t-test, 6 WT mice for each age; * p < 0.05). (P) Western blot of mouse CSF (150μL per age, pooled from ∼ 10 mice) from pre- (P20) and post-CP (P45) time points. Exposure time for Otx2 was 200-fold longer than for TTR.

Error bars, SEM.

Neither the number of calretinin-positive cells (Fig. 3D, I, K; 93.9 ± 2.3 %, p > 0.05) nor their mean intensity (Fig. 3L; Vehicle, 100 ± 1.5 vs CRE-Tat, 101.7 ± 1.5, p > 0.4) was altered, indicating a PV-cell specific effect. In addition, given that the CSF might convey Otx2 more broadly, we checked marker levels in the BLA. In this region, a decrease in Otx2, PV and PNN staining was observed as early as 4 days after CRE-Tat injection, compared to saline controls (Otx2, 65.4 ± 5.4%, p = 0.0068; PV, 66.3 ± 3.1%, p = 0.0041; WFA, 77.4 ± 1.0%, p = 0.0020; n=6 for each group). This indicates that the kinetics of Otx2 regulation from the choroid plexus may differ by brain region. Importantly, the total number of neurons identified by NeuN-staining was not modified in BLA (100.5 ± 9.1%, p > 0.4; n=6 for each group) nor that of GABA-stained cells in V1 or the intensity of GABA staining (Fig. 3E, J, K, L; Cell number: 100.3 ± 1.3%, p > 0.4; Intensity: Vehicle, 100 ± 2.1 vs CRE-Tat, 100.7 ± 2.1, p > 0.4; n=8 for each group), thus discounting the possibility of cell death in both regions.

The decrease of Otx2 in layers III/IV of the visual cortex after recombination in the adult choroid plexus later than P90 is illustrated (Western blot) and quantified in Figure 3M. Comparison with the time course of Otx2 accumulation in layer III/IV containing PV-cells (Fig. 3N) demonstrates that following recombination the level of Otx2 is similar to that observed at CP opening (P21). Interestingly, Otx2 mRNA expression (RT-PCR) by the choroid plexus increased only slightly between P7 and P90 (Fig. 3O); a similar pattern was seen for total Otx2 protein (data not shown). This constant expression before and after the visual CP was also seen at the level of surface associated Otx2 protein (4.6 % pre-CP and 5.4 % post-CP), as well as for Otx2 in the CSF (Fig. 3P), even though the low amounts of protein in the CSF (see above) make quantification difficult. Taken together, these results suggest that the amount of Otx2 in PV-cells is not regulated by the levels expressed by the choroid plexus and released into the CSF, but rather by the ability of the cells to capture the homeoprotein. This conclusion is consistent with previous results demonstrating that PNN assembly regulates Otx2 internalization during development and in the adult (Beurdeley et al., 2012; Miyata et al., 2012; Sugiyama et al., 2008).

Knocking-down Otx2 in the choroid plexus restores plasticity in visual cortex

Down-regulation of PV expression and PNN structures by direct blockade of Otx2 transfer into visual cortex induces a ‘juvenile’ stage permissive for plasticity (Beurdeley et al., 2012). To test the functional impact of choroid-derived Otx2, we used visually-evoked potential (VEP) recordings to measure visual acuity. In adult (>P90) wild-type mice, a short-term monocular deprivation (STMD; 4 days) fails to impair vision (Bavelier et al., 2010; Prusky and Douglas, 2003). In strong contrast, STMD imposed 2 weeks after a single CRE-Tat injection into the ventricles of adult Otx2flox/flox mice significantly decreased visual acuity, indicating a reactivation of cortical plasticity (Fig. 4A, B & S1; Vehicle: 0.49 ± 0.02 vs CRE-Tat: 0.31 ± 0.03 cycles / deg, p = 0.004).

Figure 4.

Figure 4

Open in a new tab

Choroid Otx2 disruption reactivates plasticity in adult visual cortex.

(A) Schematic of adult (>P90) visual cortical plasticity paradigm by short-term (4 day) monocular deprivation (STMD).

(B) Visual acuity in untreated adult wild-type or Otx2flox/flox mice injected icv with either vehicle or CRE-Tat protein. Two weeks post-injection, Otx2flox/flox mice underwent STMD (or not) just before VEP recording. Horizontal bar, mean value for each group (paired t test; 3-5 mice per proup; ** p<0.01).

(C) Schematic of amblyopia recovery paradigm. Amblyopia is induced during the CP via a long-term (14 days) monocular deprivation (LTMD). CRE-Tat injections are made 4 weeks after eye suture removal. VEP acuity is then measured in adulthood (>P90).

(D) Visual acuity recovers significantly in adult Otx2flox/flox mice injected icv with CRE-Tat protein. (paired t test; 4 mice per group; *** p<0.001).

(E) A two-threshold model for Otx2 regulation of cortical plasticity. Early sensory experience initiates the accumulation of Otx2 in PV-cells to promote their functional maturation and plasticity onset. As the CP proceeds, PNNs condense tightly around these PV-cells causing higher levels of Otx2 to accumulate and to structurally stabilize the local circuit.

See also Figure S1.

The therapeutic value of reducing choroid-derived Otx2 was further confirmed by monitoring recovery from amblyopia induced earlier in life. Ventricular CRE-Tat injection in adult Otx2flox/flox mice during the recovery period rescued visual acuity from an initial long-term deprivation spanning the CP (Fig. 4C,D & S1; Vehicle: 0.26 ± 0.01 vs CRE-Tat: 0.38 ± 0.01 cycles / deg, p = 0.0005). These results are comparable to those obtained after direct blockade of Otx2 transfer within visual cortex (Beurdeley et al., 2012). Specific interactions between a sugar-binding motif in Otx2 and PNNs maintains a metastable physiological state of PV-cells (Beurdeley et al., 2012; Miyata et al., 2012). Taken together, plasticity may be activated at a low Otx2 threshold and then actively repressed in adulthood by the continuous transfer of choroid-derived Otx2 into mature PV-cells after a second threshold has been passed. This model (Fig. 4E) is discussed below.

Discussion

In a previous study, we have shown that one allele of Otx2 is not enough to induce plasticity in V1 (Sugiyama et al., 2008). Conversely, here we demonstrate that reducing adult Otx2 content by only 32% is enough to reopen plasticity. This suggests a physiological range for Otx2 action defined by two thresholds (Fig. 4E), one to open and another one to close plasticity. This is reminiscent of a ‘French flag’ model of maturation similar to that proposed by Lewis Wolpert (Wolpert, 1969), here a temporal model as the morphogen concentration increases with time within PV-cells. Importantly, the second threshold is ‘reversible’ as decreasing Otx2 concentration engages the plastic regime (this study; (Beurdeley et al., 2012)).

The model also stipulates that early sensory experience (sensory input) initiates expression of Otx2 binding sites on nascent PNNs (Beurdeley et al., 2012; Miyata et al., 2012), facilitating Otx2 import and opening a period of plasticity. Subsequent Otx2 internalization increases intracellular Otx2 and PV-cell maturation reflected by further PV expression and PNN assembly (Sugiyama et al., 2008) until a second threshold is reached that closes plasticity. In support of the hypothesis is that direct infusion of Otx2 before CP opening opens CP ahead of time and also closes it ahead of time (Sugiyama et al., 2008). Moreover, the opening of plasticity (first threshold) does not take place in dark-reared animals, where the Otx2 and PV levels remain low in PV-cells. Instead, PV-cell maturation in dark-reared animals is rescued by direct Otx2 infusion leading to its specific accumulation in PV-cells and CP opening (Sugiyama et al., 2008).

Together, this indicates that the ability of PV-cells to capture Otx2 after initial PNN assembly may be the key event that regulates plasticity independently of age. This is further supported by the demonstration that the knock-out of specific sulfotranferases that modify the PNN structure interferes with Otx2 internalization and CP closure (Miyata et al., 2012) and that the permanent transfer of Otx2 through PNN recognition is necessary to repress plasticity in the adult (Beurdeley et al., 2012). The finding that hydrolysis of chondroitin sulfates reopens plasticity in the adult rat visual cortex may thus be explained, in part, by the persistent role of PNNs in Otx2 transport into PV-cells (Pizzorusso et al., 2002). Overall, one can propose that plasticity during development and in the adult is regulated by Otx2 import and that PNNs are the gatekeepers of Otx2.

The source of Otx2 then becomes an important point to consider. Blocking Otx2 transport in the retina (between bipolar cells and retinal ganglion cells) or in the cortex (by directly infusing anti-Otx2 antibodies) retards the activity-dependent opening of CP in the visual cortex (Sugiyama et al., 2008). The activity-dependent CP opening may thus require the transport of Otx2 along the visual pathway and ensuing initial maturation of PV-cells marked by PNN assembly. Accordingly, we showed that Otx2 injected in the eye can travel along this pathway and terminates in PV-cells (Sugiyama et al., 2008; Sugiyama et al., 2009). It is also plausible that neural activity per se regulates PNN assembly through another mechanism and that Otx2 from other sources is internalized by the maturing PV-cells.

The two possibilities are not mutually exclusive and the most important point is that the first threshold (Fig. 4E) is activity-dependent. The observation that Otx2 is present in PV-cells outside the visual cortex (Fig. 1) favors amore general source, supported here by the choroid plexus. The activity-dependent opening of plasticity based on PNN assembly and Otx2 capture might explain why CPs do not open uniformly throughout the cortex even though the choroid plexus provides a continuous supply of Otx2. Salient patterns of neural input emerge at different times as the various peripheral sense organs come on-line. The same scheme also explain show a general source, thanks to mature, fully assembled PNNs, participates in maintaining a non-plastic state, allowing for the possibility to reopen plasticity in adulthood (Beurdeley et al., 2012; Pizzorusso et al., 2002).

Thus, one striking implication is that the choroid plexus, via the CSF, may be a general regulator of CP plasticity, including the amygdala (Gogolla et al., 2009), auditory (Barkat et al., 2011) or prefrontal cortex (Belforte et al., 2010; Yang et al., 2012) – structures with shifted or shorter critical periods than V1. An important Otx2 target may be the enriched ATP metabolism in these fast-spiking cells (Plessy et al., 2008). Indeed, translation of nuclear-encoded mitochondrial mRNAs commonly follows homeoprotein capture by dopaminergic neurons or retinal ganglion cell growth cones (Alvarez-Fischer et al., 2011; Stettler et al., 2012; Yoon et al., 2012). Correspondingly, PV-cells are highly vulnerable to oxidative stress across a number of cognitive disorders (Do et al., 2009), which may be protected by PNNs (Cabungcal et al., 2013).

The choroid plexus then offers a novel route to access brain plasticity through the peripheral blood supply (Johanson et al., 2008). For example, cell-permeable peptides injected into the retro-orbital sinus can efficiently deliver reagents (eg. morpholinos, siRNA) to transiently knock-down Otx2 synthesis in the choroid plexus (see Fig. S2). The differential kinetics of Otx2 regulation across brain regions might then be paired effectively with therapeutic strategies for neurodevelopmental disorders more broadly.

Material and Methods

Animals

Conventionally raised C57Bl/6 mice (12:12 hours light:dark cycle) were purchased from Janvier (Le Genest-Saint-Isle, France). ROSA26R reporter mice were obtained from the Jackson Laboratory. Otx2flox/flox mice were generated and kindly donated by Dr. T. Lamonerie (IBDC, Nice, France) (Fossat et al., 2006) and S Aizawa (RIKEN CDB, Kobe, Japan) (Nishida et al., 2003). All procedures were designed to minimize animal suffering and carried out in accordance with the recommendations of the European Economic Community (86/609/EEC), the French National Committee (87/848) for the use of laboratory animals and the IACUC committee of Boston Children's Hospital.

Cell surface biotinylation

Adult mouse choroid plexus from the lateral and fourth ventricles were rapidly extracted in PBS and immediately processed for biotinylation with EZlink-SulfoNHS-Biotin (Pierce #21326) in PBS for 30 min at RT with gentle agitation. Following several washes, tissues were disrupted by sonication in the presence of protease inhibitors and nucleases. After clarification, the lysate was incubated with magnetic streptavidin beads (Dynabeads, #65001) for 20 min at 4°C. Following intensive washing, beads were directly resuspended in SDS-PAGE loading buffer and eluted biotinylated proteins were processed for Western Blot detection with the indicated antibodies.

CSF sampling

The CSF sampling procedure was adapted from described methods (Liu and Duff, 2008; Morrey et al., 2008). Briefly, anesthetized mice were mounted on a stereotaxic frame, the dura covering the cisterna magna was gently exposed and the surrounding area thoroughly cleaned using sterile saline-soaked cotton swabs. CSF was mechanically aspirated through a 30-gauge needle connected to a sterile polyvinylchloride catheter tube (ID: 0.69 mm, OD: 1.14 mm; Alzet) and inserted into the cisterna magna. On average, 15μL of CSF could be sampled from an adult mouse. Any samples contaminated with the slightest traces of blood were discarded. During the collection procedure, CSF samples were kept on ice, in the presence of PMSF (1mM). Prior to acetone-mediated total protein precipitation, CSF was subjected to centrifugation (for 1 min at 1000g and then 20 min at 17000g) to remove cells and cell fragments. The protein pellet was finally resuspended in SDS-PAGE loading buffer and processed for Western Blot detection.

Protein production

The coding sequence of CRE-Tat protein was inserted in a derivative of pSCherry2 (Eurogentec) and expressed in SE1 cells grown in MagicMedia Medium (Invitrogen) according to manufacturer's instructions (details of the constructs available on request). Briefly, cells were grown for 18h at 37°C and then harvested by centrifugation. Pellets were frozen at -80°C. Bacteria were resuspended in buffer A (300 mM NaCl, 10 mM Phosphate buffer, 30 mM imidazole, pH 7.5 with protease inhibitors; RocheDiagnostics) and disrupted by sonication (Bioruptor, Diagenode). The lysate was clarified by centrifugation (13000 g, 20 min, 4°C) and passed through a 0.45 μm filter. The protein was purified by two successive steps of affinity chromatography, first on Nickel column and second on heparin column following removal of the polyhistidine tag by preScisson proteolytic cleavage. The recombinant protein was eluted on FPLC (AKTA, GE Healthcare), with a linear gradient of NaCl (ranging from 0.3 to 2 M), in 10 mM Phosphate buffer, pH 7.5. Fractions were quantified and analysed on SDS-PAGE and stored at -20°C directly in the elution buffer.

Protein injection

CRE-Tat recombinant protein (or vehicle) was injected bilaterally into the lateral ventricles of adult mice, using a 10 μL Hamilton syringe coupled to an electronic injection device, at a rate of 0.2 μL /min. Coordinates for injections were (with respect to bregma): -0.58 mm on the A/P axis; +/- 1.28 mm laterally; 2 mm in depth. 5 days after injection, injected ROSA26R reporter mice were perfused transcardially with phosphate buffered saline (PBS) followed by 4% para-formaldehyde (PFA) prepared in PBS. After 4 hours of post-fixation in 4% PFA, brains were washed in PBS. Coronal brain sections (50 μm) were cut on a vibratome (Microm) and processed for X-Gal staining. Otx2flox/flox mice were sacrificed two weeks after injection. Following intracardiac PBS perfusion, the retina, choroid plexus (pooled lateral and 4th ventricles), dLGN and superior colliculus were rapidly extracted in PBS, frozen on dry ice and stored at -80°C. Cerebral cortices were left intact and further processed for immunostaining. To that end, tissues were fixed for 6 hours in 4% PFA, washed with PBS and sliced on a vibratome. Alternatively, posterior cortex layer IV was dissected under a dissection microscope, frozen on dry ice and stored at -80°C prior to total protein lysate preparation and Western Blot detection.

DNA, RNA and protein extraction

Using the All Prep DNA/RNA Mini Kit (Qiagen), genomic DNA, total RNA and proteins could be simultaneously recovered from the same biological samples. Frozen choroid plexus, retina, dLGN and superior colliculus were processed following manufacturer's instructions.

PCR

PCR was used to confirm the efficiency of CRE-induced recombination at the level of genomic DNA. To that end, we used C and E primers as previously described (Fossat et al., 2006). As these primers hybridize on either side of the flox cassettes, excised and wild type alleles could be distinguished by the size of the amplified fragments.

Quantitative RT-PCR

First-strand cDNA was synthesized using 100 ng of total mRNA, a blend of Omniscript and Sensiscript Reverse Transcriptases and a mix of oligo-dT and random primers (QuantiTect Rev. Transcription Kit, Qiagen). For real time PCR, samples were analyzed in triplicate with a LightCycler 480 II (Roche). Gene to GAPDH ratios were determined using the 2-ΔΔCt method.

Western Blot

Proteins from choroid plexus, retina, dLGN, superior colliculus, cortical layer IV and CSF samples were separated on NuPAGE 4–12% Bis-Tris precasted gels (Invitrogen) and transferred onto PVDF membrane. Primary antibodies were incubated overnight at 4°C and HRP-coupled secondary antibodies for 1 hour at room temperature. The following antibodies were used: anti-Otx2 (rat polyclonal, 1/2000)(Sugiyama et al., 2008), anti α-tubulin (mouse monoclonal, clone B-5-1-2, 1/75000, Sigma-Aldrich), anti RhoA (mouse monoclonal, clone 26CA, 1/500, Santa Cruz Biotechnology), anti-TTR (rabbit polyclonal, 1/2000, Interchim) and anti-TrkB (rabbit monoclonal, clone 80E3, 1/1000, CST). Corresponding HRP-coupled secondary antibodies were purchased from GE Healthcare (1/4000 – 1/10000).

In situ hybridization and immunostaining

In situ hybridization was performed as described (Sugiyama et al., 2008). Briefly, 50 μm floating sections were hybridized overnight at 70°C with a Digoxigenin (DIG)-labeled RNA probe for Otx2 mRNA. The binding of the probe was revealed using an Alkaline Phosphatase (AP)-coupled anti-DIG antibody (sheep polyclonal, 1/1000, Roche Applied Science). For immunostaining, floating sections were incubated with primary antibodies overnight at 4°C, intensively washed at room temperature and further incubated with corresponding Alexa-Fluor-conjugated secondary antibodies overnight at 4°C. The following primary antibodies were used: anti-Otx2 (rat polyclonal, 1/200) (Sugiyama et al., 2008), anti-PV (mouse monoclonal, clone PARV-19, 1/400, Sigma-Aldrich), anti-Calretinin (rabbit serum, 1/2000, Swant) and anti-GABA (rabbit polyclonal, 1/1000, Sigma-Aldrich). Secondary antibodies were used as follows: anti-rat Alexa-Fluor 561 (1/5000, Molecular Probes), anti-Mouse Alexa-Fluor 488 (1/5000, Molecular Probes) and anti-Rabbit Alexa-Fluor 633 (1/5000, Molecular Probes). Biotinylated-WFA was purchased from Sigma-Aldrich (1/100) and was detected by means of Alexa-Fluor 633-conjugated streptavidin (Molecular probes). Stained sections were mounted in Fluoromount medium (Southern Biotech) and images were acquired using a Leica SP5 confocal microscope.

Visual evoked potentials (VEPs)

Electrophysiological recordings were performed under nembutal/chlorprothixene anesthesia using standard techniques (Beurdeley et al., 2012). Transient VEPs in response to abrupt contrast reversal (100%, 1 Hz) over a range of spatial frequencies (0.01 – 0.6 cycles / degree) were band-pass-filtered (0.1–100 Hz), amplified, and fed to a computer for analysis. In brief, at least 20 events were averaged in synchrony with the stimulus contrast reversal. Visual acuity was obtained by extrapolation to zero amplitude after correction of noise level. Monocular deprivation consisted of eyelid suture under isoflurane anesthesia; in some cases, the suture was reopened to test for recovery.

Supplementary Material

supp figures
supp inventory
supp legends

Acknowledgments

We thank Prof. M. Volovitch for useful discussions. In addition to institutional support, this study was funded by grants to A.J. and A.P. from Agence Nationale pour la Recherche (ANR-10-BLAN-141-01 & ANR-11-BLAN-069467), Global Research Laboratory Program (2009-00424) from the Korean Ministry of Education, Science and Technology and Region Ile de France. T.K.H. was supported by NIH (1 DP1 OD 003699), J.S. was a College de France Post-Doctoral Fellow, and H.H.C.L. was a Croucher Foundation (Hong Kong) Fellow.

References

  1. Alonso MI, Martin C, Carnicero E, Bueno D, Gato A. Cerebrospinal fluid control of neurogenesis induced by retinoic acid during early brain development. Dev Dyn. 2011;240:1650–1659. doi: 10.1002/dvdy.22657. [DOI] [PubMed] [Google Scholar]
  2. Alvarez-Fischer D, Fuchs J, Castagner F, Stettler O, Massiani-Beaudoin O, Moya KL, Bouillot C, Oertel WH, Lombes A, Faigle W, et al. Engrailed protects mouse midbrain dopaminergic neurons against mitochondrial complex I insults. Nat Neurosci. 2011;14:1260–1266. doi: 10.1038/nn.2916. [DOI] [PubMed] [Google Scholar]
  3. Barkat TR, Polley DB, Hensch TK. A critical period for auditory thalamocortical connectivity. Nat Neurosci. 2011;14:1189–1194. doi: 10.1038/nn.2882. [DOI] [PMC free article] [PubMed] [Google Scholar]
  4. Bavelier D, Levi DM, Li RW, Dan Y, Hensch TK. Removing brakes on adult brain plasticity: from molecular to behavioral interventions. J Neurosci. 2010;30:14964–14971. doi: 10.1523/JNEUROSCI.4812-10.2010. [DOI] [PMC free article] [PubMed] [Google Scholar]
  5. Belforte JE, Zsiros V, Sklar ER, Jiang Z, Yu G, Li Y, Quinlan EM, Nakazawa K. Postnatal NMDA receptor ablation in corticolimbic interneurons confers schizophrenia-like phenotypes. Nat Neurosci. 2010;13:76–83. doi: 10.1038/nn.2447. [DOI] [PMC free article] [PubMed] [Google Scholar]
  6. Beurdeley M, Spatazza J, Lee H, Sugiyama S, Bernard C, Di Nardo AA, Hensch TK, Prochiantz A. Otx2 binding to perineuronal nets persistently regulates plasticity in the mature visual cortex. J Neurosci. 2012;32:9429–9437. doi: 10.1523/JNEUROSCI.0394-12.2012. [DOI] [PMC free article] [PubMed] [Google Scholar]
  7. Cabungcal JH, Steullet P, Kraftsik R, Cuenod M, Do KQ. Early-Life Insults Impair Parvalbumin Interneurons via Oxidative Stress: Reversal by N-Acetylcysteine. Biol Psychiatry. 2013;73:574–582. doi: 10.1016/j.biopsych.2012.09.020. [DOI] [PubMed] [Google Scholar]
  8. Di Lullo E, Haton C, Le Poupon C, Volovitch M, Joliot A, Thomas JL, Prochiantz A. Paracrine Pax6 activity regulates oligodendrocyte precursor cell migration in the chick embryonic neural tube. Development. 2011;138:4991–5001. doi: 10.1242/dev.066282. [DOI] [PubMed] [Google Scholar]
  9. Do KQ, Cabungcal JH, Frank A, Steullet P, Cuenod M. Redox dysregulation, neurodevelopment, and schizophrenia. Curr Opin Neurobiol. 2009;19:220–230. doi: 10.1016/j.conb.2009.05.001. [DOI] [PubMed] [Google Scholar]
  10. Elbert T, Pantev C, Wienbruch C, Rockstroh B, Taub E. Increased cortical representation of the fingers of the left hand in string players. Science. 1995;270:305–307. doi: 10.1126/science.270.5234.305. [DOI] [PubMed] [Google Scholar]
  11. Fossat N, Chatelain G, Brun G, Lamonerie T. Temporal and spatial delineation of mouse Otx2 functions by conditional self-knockout. EMBO Rep. 2006;7:824–830. doi: 10.1038/sj.embor.7400751. [DOI] [PMC free article] [PubMed] [Google Scholar]
  12. Gogolla N, Caroni P, Luthi A, Herry C. Perineuronal nets protect fear memories from erasure. Science. 2009;325:1258–1261. doi: 10.1126/science.1174146. [DOI] [PubMed] [Google Scholar]
  13. Hensch TK. Critical period regulation. Annu Rev Neurosci. 2004;27:549–579. doi: 10.1146/annurev.neuro.27.070203.144327. [DOI] [PubMed] [Google Scholar]
  14. Hensch TK. Critical period plasticity in local cortical circuits. Nat Rev Neurosci. 2005;6:877–888. doi: 10.1038/nrn1787. [DOI] [PubMed] [Google Scholar]
  15. Huang X, Liu J, Ketova T, Fleming JT, Grover VK, Cooper MK, Litingtung Y, Chiang C. Transventricular delivery of Sonic hedgehog is essential to cerebellar ventricular zone development. Proc Natl Acad Sci U S A. 2010;107:8422–8427. doi: 10.1073/pnas.0911838107. [DOI] [PMC free article] [PubMed] [Google Scholar]
  16. Johanson CE, Duncan JA, 3rd, Klinge PM, Brinker T, Stopa EG, Silverberg GD. Multiplicity of cerebrospinal fluid functions: New challenges in health and disease. Cerebrospinal Fluid Res. 2008;5:10. doi: 10.1186/1743-8454-5-10. [DOI] [PMC free article] [PubMed] [Google Scholar]
  17. Johansson PA, Irmler M, Acampora D, Beckers J, Simeone A, Gotz M. The transcription factor Otx2 regulates choroid plexus development and function. Development. 2013;140:1055–1066. doi: 10.1242/dev.090860. [DOI] [PubMed] [Google Scholar]
  18. Koike C, Nishida A, Ueno S, Saito H, Sanuki R, Sato S, Furukawa A, Aizawa S, Matsuo I, Suzuki N, et al. Functional roles of Otx2 transcription factor in postnatal mouse retinal development. Mol Cell Biol. 2007;27:8318–8329. doi: 10.1128/MCB.01209-07. [DOI] [PMC free article] [PubMed] [Google Scholar]
  19. Kuhl PK, Williams KA, Lacerda F, Stevens KN, Lindblom B. Linguistic experience alters phonetic perception in infants by 6 months of age. Science. 1992;255:606–608. doi: 10.1126/science.1736364. [DOI] [PubMed] [Google Scholar]
  20. Lehtinen MK, Zappaterra MW, Chen X, Yang YJ, Hill AD, Lun M, Maynard T, Gonzalez D, Kim S, Ye P, et al. The cerebrospinal fluid provides a proliferative niche for neural progenitor cells. Neuron. 2011;69:893–905. doi: 10.1016/j.neuron.2011.01.023. [DOI] [PMC free article] [PubMed] [Google Scholar]
  21. Liu L, Duff K. A technique for serial collection of cerebrospinal fluid from the cisterna magna in mouse. J Vis Exp. 2008 doi: 10.3791/960. [DOI] [PMC free article] [PubMed] [Google Scholar]
  22. Martin C, Bueno D, Alonso MI, Moro JA, Callejo S, Parada C, Martin P, Carnicero E, Gato A. FGF2 plays a key role in embryonic cerebrospinal fluid trophic properties over chick embryo neuroepithelial stem cells. Dev Biol. 2006;297:402–416. doi: 10.1016/j.ydbio.2006.05.010. [DOI] [PubMed] [Google Scholar]
  23. Maurer D, Hensch TK. Amblyopia: background to the special issue on stroke recovery. Dev Psychobiol. 2012;54:224–238. doi: 10.1002/dev.21022. [DOI] [PubMed] [Google Scholar]
  24. Miyata S, Komatsu Y, Yoshimura Y, Taya C, Kitagawa H. Persistent cortical plasticity by upregulation of chondroitin 6-sulfation. Nat Neurosci. 2012;15:414–422. doi: 10.1038/nn.3023. [DOI] [PubMed] [Google Scholar]
  25. Morrey JD, Olsen AL, Siddharthan V, Motter NE, Wang H, Taro BS, Chen D, Ruffner D, Hall JO. Increased blood-brain barrier permeability is not a primary determinant for lethality of West Nile virus infection in rodents. J Gen Virol. 2008;89:467–473. doi: 10.1099/vir.0.83345-0. [DOI] [PubMed] [Google Scholar]
  26. Nelson CA, 3rd, Zeanah CH, Fox NA, Marshall PJ, Smyke AT, Guthrie D. Cognitive recovery in socially deprived young children: the Bucharest Early Intervention Project. Science. 2007;318:1937–1940. doi: 10.1126/science.1143921. [DOI] [PubMed] [Google Scholar]
  27. Nishida A, Furukawa A, Koike C, Tano Y, Aizawa S, Matsuo I, Furukawa T. Otx2 homeobox gene controls retinal photoreceptor cell fate and pineal gland development. Nat Neurosci. 2003;6:1255–1263. doi: 10.1038/nn1155. [DOI] [PubMed] [Google Scholar]
  28. Nothias F, Fishell G, Ruiz i Altaba A. Cooperation of intrinsic and extrinsic signals in the elaboration of regional identity in the posterior cerebral cortex. Curr Biol. 1998;8:459–462. doi: 10.1016/s0960-9822(98)70189-7. [DOI] [PubMed] [Google Scholar]
  29. Pizzorusso T, Medini P, Berardi N, Chierzi S, Fawcett JW, Maffei L. Reactivation of ocular dominance plasticity in the adult visual cortex. Science. 2002;298:1248–1251. doi: 10.1126/science.1072699. [DOI] [PubMed] [Google Scholar]
  30. Plessy C, Fagiolini M, Wagatsuma A, Harasawa N, Kuji T, Asaka-Oba A, Kanzaki Y, Fujishima S, Waki K, Nakahara H, et al. A resource for transcriptomic analysis in the mouse brain. PLoS One. 2008;3:e3012. doi: 10.1371/journal.pone.0003012. [DOI] [PMC free article] [PubMed] [Google Scholar]
  31. Prusky GT, Douglas RM. Developmental plasticity of mouse visual acuity. Eur J Neurosci. 2003;17:167–173. doi: 10.1046/j.1460-9568.2003.02420.x. [DOI] [PubMed] [Google Scholar]
  32. Puelles E, Annino A, Tuorto F, Usiello A, Acampora D, Czerny T, Brodski C, Ang SL, Wurst W, Simeone A. Otx2 regulates the extent, identity and fate of neuronal progenitor domains in the ventral midbrain. Development. 2004;131:2037–2048. doi: 10.1242/dev.01107. [DOI] [PubMed] [Google Scholar]
  33. Rath MF, Morin F, Shi Q, Klein DC, Moller M. Ontogenetic expression of the Otx2 and Crx homeobox genes in the retina of the rat. Exp Eye Res. 2007;85:65–73. doi: 10.1016/j.exer.2007.02.016. [DOI] [PubMed] [Google Scholar]
  34. Rath MF, Munoz E, Ganguly S, Morin F, Shi Q, Klein DC, Moller M. Expression of the Otx2 homeobox gene in the developing mammalian brain: embryonic and adult expression in the pineal gland. J Neurochem. 2006;97:556–566. doi: 10.1111/j.1471-4159.2006.03773.x. [DOI] [PubMed] [Google Scholar]
  35. Sawamoto K, Wichterle H, Gonzalez-Perez O, Cholfin JA, Yamada M, Spassky N, Murcia NS, Garcia-Verdugo JM, Marin O, Rubenstein JL, et al. New neurons follow the flow of cerebrospinal fluid in the adult brain. Science. 2006;311:629–632. doi: 10.1126/science.1119133. [DOI] [PubMed] [Google Scholar]
  36. Simeone A, Acampora D, Mallamaci A, Stornaiuolo A, D'Apice MR, Nigro V, Boncinelli E. A vertebrate gene related to orthodenticle contains a homeodomain of the bicoid class and demarcates anterior neuroectoderm in the gastrulating mouse embryo. EMBO J. 1993;12:2735–2747. doi: 10.1002/j.1460-2075.1993.tb05935.x. [DOI] [PMC free article] [PubMed] [Google Scholar]
  37. Soriano P. Generalized lacZ expression with the ROSA26 Cre reporter strain. Nat Genet. 1999;21:70–71. doi: 10.1038/5007. [DOI] [PubMed] [Google Scholar]
  38. Stettler O, Joshi RL, Wizenmann A, Reingruber J, Holcman D, Bouillot C, Castagner F, Prochiantz A, Moya KL. Engrailed homeoprotein recruits the adenosine A1 receptor to potentiate ephrin A5 function in retinal growth cones. Development. 2012;139:215–224. doi: 10.1242/dev.063875. [DOI] [PubMed] [Google Scholar]
  39. Sugiyama S, Di Nardo AA, Aizawa S, Matsuo I, Volovitch M, Prochiantz A, Hensch TK. Experience-dependent transfer of Otx2 homeoprotein into the visual cortex activates postnatal plasticity. Cell. 2008;134:508–520. doi: 10.1016/j.cell.2008.05.054. [DOI] [PubMed] [Google Scholar]
  40. Sugiyama S, Prochiantz A, Hensch TK. From brain formation to plasticity: insights on Otx2 homeoprotein. Dev Growth Differ. 2009;51:369–377. doi: 10.1111/j.1440-169X.2009.01093.x. [DOI] [PubMed] [Google Scholar]
  41. Timmusk T, Mudo G, Metsis M, Belluardo N. Expression of mRNAs for neurotrophins and their receptors in the rat choroid plexus and dura mater. Neuroreport. 1995;6:1997–2000. doi: 10.1097/00001756-199510010-00011. [DOI] [PubMed] [Google Scholar]
  42. Weisner B, Roethig HJ. The concentration of prealbumin in cerebrospinal fluid (CSF), indicator of CSF circulation disorders. Eur Neurol. 1983;22:96–105. doi: 10.1159/000115544. [DOI] [PubMed] [Google Scholar]
  43. Wizenmann A, Brunet I, Lam JSY, Sonnier L, Beurdeley M, Zarbalis K, Weisenhorn-Vogt D, Weiln C, Dwivedy A, Joliot A, et al. Extracellular Engrailed participates in the topographic guidance of retinal axons in vivo. Neuron. 2009;64:355–366. doi: 10.1016/j.neuron.2009.09.018. [DOI] [PMC free article] [PubMed] [Google Scholar]
  44. Wolpert L. Positional information and the spatial pattern of cellular differentiation. J Theor Biol. 1969;25:1–47. doi: 10.1016/s0022-5193(69)80016-0. [DOI] [PubMed] [Google Scholar]
  45. Yang EJ, Lin EW, Hensch TK. Critical period for acoustic preference in mice. Proc Natl Acad Sci U S A. 2012;109(Suppl 2):17213–17220. doi: 10.1073/pnas.1200705109. [DOI] [PMC free article] [PubMed] [Google Scholar]
  46. Yoon BC, Jung H, Dwivedy A, O'Hare CM, Zivraj KH, Holt CE. Local translation of extranuclear lamin B promotes axon maintenance. Cell. 2012;148:752–764. doi: 10.1016/j.cell.2011.11.064. [DOI] [PMC free article] [PubMed] [Google Scholar]

Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Supplementary Materials

supp figures
NIHMS606809-supplement-supp_figures.pdf (353.4KB, pdf)
supp inventory
NIHMS606809-supplement-supp_inventory.docx (44.8KB, docx)
supp legends
NIHMS606809-supplement-supp_legends.docx (117.2KB, docx)

RESOURCES